Helicobacter pylori actively suppresses innate immune nucleic acid receptors

ABSTRACT Chronic mucosal pathogens have evolved multiple strategies to manipulate the host immune response; consequently, microbes contribute to the development of >2 million cases of cancer/year. Gastric adenocarcinoma is the fourth leading cause of cancer-related death and Helicobacter pylori confers the highest risk for this disease. Gastric innate immune effectors can either eliminate bacteria or mobilize adaptive immune responses including Toll-like receptors (TLRs), and cytosolic DNA sensor/adaptor proteins (e.g., stimulator of interferon genes, STING). The H. pylori strain-specific cag type IV secretion system (T4SS) augments gastric cancer risk and translocates DNA into epithelial cells where it activates the microbial DNA sensor TLR9 and suppresses injury in vivo; however, the ability of H. pylori to suppress additional nucleic acid PRRs within the context of chronic gastric inflammation and injury remains undefined. In this study, in vitro and ex vivo experiments identified a novel mechanism through which H. pylori actively suppresses STING and RIG-I signaling via downregulation of IRF3 activation. In vivo, the use of genetically deficient mice revealed that Th17 inflammatory responses are heightened following H. pylori infection within the context of Sting deficiency in conjunction with increased expression of a known host immune regulator, Trim30a. This novel mechanism of immune suppression by H. pylori is likely a critical component of a finely tuned rheostat that not only regulates the initial innate immune response, but also drives chronic gastric inflammation and injury.


Introduction
Mucosal pathogens have evolved multiple strategies to manipulate the host immune response; 1,2 consequently, microbes contribute to the development of greater than 2 million cases of cancer/year. 3 Gastric adenocarcinoma is the fourth leading cause of cancerrelated death [4][5][6] and chronic infection with Helicobacter pylori confers the highest known risk for this disease. 3,6 Initial components of the innate immune system encountered by H. pylori include epithelial cells, macrophages, and dendritic cells, and interactions between H. pylori and these constituents dysregulate signaling pathways that influence oncogenesis. 7,8 Epithelial cells express effectors that can either eliminate bacteria or mobilize adaptive immune responses; these include patternrecognition receptors (PRRs), which detect and respond to conserved microbial motifs. 9,10 Functionally distinct PRR subclasses include Nodlike receptors (NLRs), Toll-like receptors (TLRs), and cytosolic DNA sensor/adaptor proteins (e.g., stimulator of interferon genes, STING), all of which are linked to gastric cancer. 10,11 PRRs orchestrate immune responses targeting pathogens and bridge innate and adaptive immunity via recognition of pathogen-associated molecular patterns (PAMPs). 9,10 However, H. pylori harbors multiple PAMPs that function differently than the respective counterparts in other mucosal pathogens. Specifically, 1) H. pylori FlaA (the major flagellin subunit) is a noninflammatory molecule in terms of its ability to activate TLR5, 1 2) H. pylori LPS induces an attenuated TLR4-mediated response, 12, 13 3) deacetylation of peptidoglycan allows H. pylori to evade host clearance via activation of a Nod1-dependent negative feedback loop, [14][15][16][17] and 4) TLR9 suppresses the injury response to this pathogen. 18 Thus, H. pylori has evolved an array of diverse phenotypes to subvert obstacles presented by the host, which promotes long-term colonization and carcinogenesis.
Microbial-specific nucleic acids are an important subclass of PAMPs, which are rapidly detected in the cytosol of host cells. 33,34 Cyclic GMP-AMP synthase (cGAS) is a cytosolic DNA sensor, which is activated in response to double-stranded DNA in a sequenceindependent manner. Binding of DNA ligands to cGAS catalyzes the conversion of ATP and GTP into the dinucleotide 2',3'-cyclic GMP-AMP (cGAMP). cGAMP can then directly activate stimulator of interferon genes (STING), a DNA sensor/adaptor localized to the endoplasmic reticulum (ER) 33,34 and which is expressed in gastric epithelial cells. 11 Sensing of cyclic dinucleotides induces a conformational change in STING that triggers trafficking of STING complexed with TANK-binding kinase 1 (TBK1) from the ER to endosomal/lysosomal compartments. Translocated TBK1 leads to phosphorylation and activation of the transcription factor interferon regulatory factor 3 (IRF3), which is then mobilized to the nucleus to induce expression of type 1 interferons (e.g., IFNα, IFNß). 33,34 STING activation can also trigger other downstream pathways such as NF-κB 35,36 as well as autophagy, which clears DNA or pathogens from the cytosol. 37 However, certain chronic pathogens have developed strategies to evade STING-mediated immune clearance, establish infection, and induce disease. 33,34,38 Carcinogenic DNA viruses, such as, human papilloma virus (HPV) 18 and human adenovirus 5 encode the oncoproteins E7 and E1A, respectively, antagonize STING. Kaposi's sarcomaassociated herpesvirus (KSHV) and hepatitis B express IRF1, tegument protein ORF52, and viral polymerases that potently disrupt the cGAS-STING pathway. Viral poxins abrogate STING signaling by degrading cGAMP. 33,34 This wide repertoire of antagonists targeting cGAS-STING underscores the importance of evolutionary pressures that select for oncogenic pathogens that can both promote malignancy and suppress innate immunity.
The role of DNA sensing in human carcinogenesis is not fully understood but recent studies indicate that DNA sensors exert a crucial role in antitumor responses. Suppression of STING in prostate and melanoma cancer cells leads to increased tumor growth, 39,40 and poor patient survival is associated with reduced cGAS and STING expression. 11,41 In gastric cancer, STING expression is significantly decreased in tumor versus non-tumor tissue, and low levels of expression are associated with reduced survival. 11 In models of inflammation with premalignant potential (e.g., chronic pancreatitis), inhibition of STING worsens disease via up-regulation of IL-17A, 42 which promotes inflammation-induced malignancies including pancreatic cancer, colitis-associated carcinoma, and skin cancer. [43][44][45] We previously demonstrated that translocated H. pylori DNA can activate the microbial DNA sensor TLR9 in vitro and that TLR9 suppresses H. pyloriinduced injury in vivo; however, the ability of H. pylori to suppress additional nucleic acid PRRs within the context of gastric inflammation and injury has not been fully investigated. Therefore, the goal of this study was to elucidate the effects of H. pylori on STING signaling, and, using a Sting-deficient mouse model, delineate the role of STING in the context of gastric inflammation and injury. Our findings identify a novel mechanism through which H. pylori actively suppresses STING-associated signaling in host cells via induction of an induced host effector, Trim30a. These pathways may contribute to the ability of H. pylori to dampen the innate immune response and ultimately promote chronic gastric inflammation and injury, which may heighten the subsequent risk for gastric carcinogenesis.

Results
We first sought to directly assess the effects of H. pylori on STING signaling, by utilizing HEK293 cells transfected with a STING-specific reporter. While levels of STING activation increased 17-fold in cells cocultured with 2ʹ3'-cGAMP (a STING agonist), activation levels in cells co-cultured with the wild-type cag + H. pylori strain J166 were no different than uninfected controls (Figure 1(a)). Because certain chronic pathogens have been shown to exert a suppressive effect on STING signaling and to facilitate long term survival, we next simultaneously co-cultured or pre-incubated cells with H. pylori prior to the addition of 2ʹ3'-cGAMP. H. pylori significantly reduced STING agonist-mediated activation by 50% in both condition (Figure 1(a)). STING suppression by H. pylori was dose- (Figure 1(b)) and time-dependent (Figure 1(c)) with minimal effects on cell viability out to 16-24 hours (Supplemental Figure 1). Previous work defining the ability of H. pylori to activate the innate immune DNA sensor TLR9 utilized strain J166. 31 Therefore, to determine if STING suppression was strain-specific, we repeated these studies using additional isolates and demonstrated that H. pylori strain Figure 1. STING activation is reduced by H. pylori in vitro. STING+ or parental cells were challenged with PBS alone (UI), 2ʹ3'-cGAMP and/or (a) wild-type (wt) cag + H. pylori strain J166, (b) H. pylori strain J166 at varying MOIs, (c) increasing pre-incubation times, or (d) cag + H. pylori strains J166, G27, PMSS1, 7.13, or B128 at MOI 100:1 for 24 hours. STING activation was assessed and is shown as fold STING activation relative to uninfected control. Experiments were performed in triplicate, and samples were run in duplicate within each experiment. ANOVA with Bonferroni correction was used to determine statistical significance among groups. (e) STING+ or parental cells were challenged with PBS alone (UI), 2ʹ3'-cGAMP and/or H. pylori strain J166 for 6 and 24 hours and were then assessed for STING expression and activation by Western blot analysis. For Western blot analysis, conditions were tested at least 3 times and student's t-tests were used to determine statistical significance between groups. *p < .05, **p < .01, ***p < .001, ****p < .0001, ns = not significant. G27, the rodent-colonizing strains PMSS1 and 7.13, and clinical isolate B128 all significantly reduced STING activation under either preincubation or coculture conditions compared to agonist alone ( Figure 1(d)). To assess whether reduced activation was attributable to reduced STING expression, we assessed STING expression and activation (pSTING) in HEK293 cells treated with the STING agonist (2ʹ3'-cGAMP) with or without H. pylori strain J166 (Figure 1(e)). These data demonstrate that HEK293 cells express similar levels of STING within all treatment groups. Similar to the STING reporter assay, activation of STING (pSTING) is significantly increased with the STING agonist (2ʹ3'-cGAMP), while H. pylori significantly reduced STING agonistmediated activation by 50% (Figure 1(e)). To determine whether 2ʹ3'-cGAMP directly altered bacterial growth or function per se, H. pylori was cultured in the absence of eukaryotic cells with varying concentrations of 2ʹ3'-cGAMP. Treatment of H. pylori with 2ʹ3'-cGAMP had no effect on H. pylori growth or the ability of H. pylori to translocate CagA into host cells (Supplemental Figure 2).
Based on the premise that translocated microbial DNA may be transcribed in host cells and subsequently activate RNA sensing molecules as previously reported in Legionella pneumophila, 46,47 the ability of H. pylori to suppress activation of RIG-I, an RNA sensor, was also investigated. In HEK293 cells transfected with a RIG-I-specific reporter, RIG-I-associated activation increased 18-fold in cells co-cultured with 3pHp-RNA, a known RIG-I agonist, compared to untreated controls (Figure 2(a)). Similar to results observed with STING activation, no RIG-I activation was observed in cells co-cultured with H. pylori alone (Figure 2(a)). However, a suppression phenotype was observed during pre-incubation with H. pylori prior to addition of 3pHp-RNA or during co-culture of H. pylori and agonist together (Figure 2(a)), and this occurred in a dose- (Figure 2(b)) and time-dependent ( Figure 2(c)) manner. Cell viability of RIG-I reporter cells was not significantly affected by the presence of H. pylori and/or agonist (Supplemental Figure 3). Mirroring the STING results, multiple H. pylori strains were able to significantly attenuate RIG-I-associated signaling in vitro (Figure 2(d)).
To determine whether the suppression phenotype for STING and RIG-I required an active interplay between H. pylori and host cells, we repeated the STING and RIG-I reporter assay experiments with wild-type H. pylori J166 that had been heatinactivated for 1 hour at 56°C. Heat-inactivation abolished the suppressive phenotype for both STING and RIG-I (Figures 3(a-b)). Further, H. pylori genomic DNA per se was unable to suppress STING-and RIG-I-associated signaling in vitro when compared to viable H. pylori (Figure 3(c-d)). These results demonstrate that only viable H. pylori can suppress STING-and RIG-I-associated signaling in vitro.
The InvivoGen HEK-Blue™ hSTING-R232 cells and HEK-Lucia™ RIG-I cells used for these assays are predominantly dependent on activation of IRF3 which, when phosphorylated, induces type I interferon (IFN) gene expression. The IFNstimulated response elements (ISRE) luciferase reporter in RIG-I reporter cells, however, can also be activated by type I IFN activation which is induced by JAK/STAT signaling through ISG54. Therefore, to determine whether the observed RIG-I phenotype was IRF3-versus JAK/STAT-dependent, we utilized the inhibitor Ruxolitinib, which inhibits downstream type I IFN signaling via the JAK-STAT pathway, leaving IRF3-dependent signaling unaffected. Ruxolitinib in the presence of H. pylori and positive agonists failed to alter suppression when compared to H. pylori and agonist alone (Figure 3(e-f)). However, when Ruxolitinib and agonist alone were co-cultured without H. pylori, RIG-I-associated signaling but not STING-associated signaling was significantly reduced compared to the agonist alone ( Figure 3(e-f)). These data suggest that H. pylori likely exerts inhibitory effects at the level of IRF in these signaling pathways. To more directly assess this, we next quantified IRF3 expression and activation (pIRF3) in HEK293 cells treated with the STING agonist (2ʹ3'-cGAMP) with or without H. pylori strain J166. These data demonstrate that the STING agonist (2ʹ3'-cGAMP) significantly increased IRF3 activation, and that this activation is significantly reduced during H. pylori coculture ( Figure 3(g)), indicating that H. pylori inhibits STING agonist-mediated IRF3 activation.
Gastroids are polarized, replenishable epithelial culture systems that can be readily generated from nontransformed gastric epithelium. 48,49 We previously developed and optimized gastroid models of H. pylori infection originating from both human and murine gastric tissues; 17,50 therefore, we capitalized on this manipulatable ex vivo system as a biologically relevant model that more faithfully recapitulates the gastric niche to extend our in vitro results using reporter systems. Primary gastric organoids generated from human patients were co-cultured for 6 or 24 hours with wild-type H. pylori strain J166, with or without the STING agonist, and protein lysates were subsequently probed for STING activation and downstream effectors of STING activation via Western blot analysis. To directly assess STING activation, we quantified STING expression and activation (pSTING) in human gastric organoids treated with the STING agonist (2ʹ3'-cGAMP) with or without H. pylori strain J166. These data demonstrate that H. pylori significantly reduced STING agonist-mediated activation, but not expression, by 3-fold, (Figure 4(a)) consistent with the results from HEK293 cells (Figure 1(e)). IRF3, an effector activated by STING, was next assessed. H. pylori-infected human organoids harbored significantly lower levels of phosphorylated IRF3, compared to uninfected controls at both 6 and 24 hours (Figure 4 (b)). The STING agonist, 2ʹ3'-cGAMP, alone induced significantly higher levels of pIRF3 at 6 hours compared to controls, but this did not occur during coculture with H. pylori; this effect diminished by 24 hours (Figure 4(b)). Concordantly, expression levels of the IRF3-dependent type I interferon stimulated genes MX1 and CXCL10 were significantly upregulated in human gastric organoids following coculture with 2ʹ3'-cGAMP (Figure 4(c)), but expression was significantly reduced in samples co-cultured with H. pylori and 2ʹ3'-cGAMP ( Figure 4(c)). These results indicate that H. pylori infection of organoids recapitulated the suppressive phenotype observed in reporter cell assays ( Figure 1). STING complexes can recruit effectors which drive autophagy-like responses, that are independent of IRF3-mediated Ifnb transcription. 51 H. pylori can also induce autophagy. Therefore, we analyzed autophagy under these conditions. All H. pylori-infected samples demonstrated a significant increase in levels of autophagy compared to uninfected samples by 24 hours when utilizing increases in the autophagosome marker LC3-II as a proxy for autophagy ( Figure 4(d)). Collectively, these results suggest that H. pylori can suppress STING signaling via IRF3 and that H. pylori-induced autophagy is independent of STING. 52,53 We next extended these findings by defining the role of STING in H. pylori-induced injury in vivo. Wild-type C57BL/6 and Sting-deficient mice (Sting −/ − ) were infected with the cag + H. pylori strain PMSS1, which harbors the ability to suppress STINGassociated signaling in vitro (Figure 1(d)). No significant differences in levels of H. pylori colonization were present between wild-type and Sting −/− mice following 8 weeks of infection ( Figure 5(a)). As expected, wildtype mice infected with H. pylori developed significantly increased levels of acute and chronic inflammation compared to uninfected controls ( Figure 5(b), Supplemental Figure 4). However, Sting genetic deficiency augmented acute ( Figure 5(b)), but not chronic inflammation (Supplemental Figure 4), compared to infected wild-type mice. Further immunophenotyping by immunohistochemistry staining revealed significantly higher levels of neutrophils among infected Sting −/− mice compared to infected wild-type mice ( Figure 5(c)), while the number of macrophages, T cells, and B cells remained unchanged regardless of Sting status (Supplemental Figure 5), suggesting that STING alters acute inflammatory events during H. pylori infection.
Decreased STING expression has been shown to be an independent and adverse predictor of overall survival in human gastric cancer patients. 11 Examination of gastric tissue from wild-type mice by immunohistochemistry revealed that levels of Sting expression among H. pylori-infected gastric tissue were significantly reduced compared to uninfected tissues ( Figure 5(d)) and, as expected, undetectable in Sting −/− mice (data not shown). Furthermore, levels of RigI were also markedly reduced in H. pyloriinfected gastric tissue compared to uninfected wildtype mice ( Figure 5(e)). These data indicate that H. pylori can not only suppress Sting activation and signaling, but can also reduce levels of Sting and RigI expression in vivo.
as fold RIG-I activation relative to uninfected control. Experiments were performed in triplicate, and samples were run in duplicate within each experiment. ANOVA with Bonferroni correction was used to determine statistical significance among groups. *p < .05, ***p < .001, ****p < .0001, ns = not significant. To identify specific effectors mediating suppression of Sting signaling by H. pylori in vivo, we next performed a discovery-based RNA-seq analysis utilizing RNA isolated from whole gastric mucosa from wild-type and Sting −/− mice. Comparisons were performed to identify Stingdependent responses at baseline and following H. pylori challenge. To first identify genes differentially expressed at baseline, datasets from uninfected wild-type and Sting −/− mice were compared ( Figure 6(a)). This comparison revealed that 336 upregulated and 325 downregulated genes were altered in a Sting-dependent manner (Supplemental Table 1). Next, to identify genes differentially expressed following H. pylori infection, datasets from uninfected versus infected wild-type mice were termed comparison 1, while uninfected versus infected Sting −/− mice were termed comparison 2 ( Figure 6(b)). Comparison 1 revealed 213 upregulated and 19 downregulated genes following H. pylori infection (Supplemental Table 2), while comparison 2 identified 840 genes differentially expressed in Sting −/− mice following H. pylori infection, with 382 upregulated and 458 downregulated genes (Supplemental Table 3). Ingenuity Pathway Analysis software was then used to harmonize the datasets to identify predicted biological functions and pathways to reveal possible mechanisms that may underpin the suppressive phenotypes (Table 1).
Pathway analysis revealed that predicted functions related to IL-17 signaling had significant activation scores (>2) in both wild-type and Sting −/− mice ( Figure 6(c)), consistent with prior data demonstrating the ability of H. pylori to induce IL-17 production and Th17 responses 17,18,[55][56][57] in conjunction with observations that inhibition of STING activation is associated with increased Th17 cell infiltration, increased production of IL-17A, and worsening inflammation in conditions such as chronic pancreatitis. 45 Therefore, to independently validate potential differences in IL-17 signaling due to Sting deficiency within the context of H. pylori infection, we examined Th17 differentiation and stabilization factors via RT-PCR from RNA isolated from gastric tissue. Differentiation and formation of Th17 cells can be driven by transcription factors such as IRF4 and cytokines such as IL-23. Inducible Costimulator (ICOS) has also been shown to be critical for the development of Th17 cells. Transcript levels of Irf4, Il23, and Icos in H. pylori-infected Sting −/− mice were significantly increased compared to infected wild-type mice ( Figure 6(d)), while no significant differences were observed in levels of Il6 and Il1b (Figure 6(d)) suggesting that, in the absence of Sting, Th17 differentiation may be driven by both IL-23 and Icos, among other factors that were not evaluated. Conversely, significantly lower levels of the Th17 inhibitors Ifng and Il12b were present among infected Sting-deficient mice ( Figure 6(d)). Transcript levels of Il17a and Il17f were also significantly increased in H. pylori-infected Sting −/− mice compared to infected wild-type mice ( Figure 6(d)). In assessing IRF3dependent type I interferon stimulated genes by RT-PCR, Mx1 and Cxcl10 were altered to a similar degree with H. pylori infection in both wild-type and Sting −/− mice ( Figure 6(e)), which validate data obtained from RNA-seq.
We next sought to recapitulate our human ex vivo findings ( Figure 4) demonstrating that H. pylori infection can selectively downregulate phosphorylated-IRF3-dependent pathways in gastric epithelial cells and further delineate the role of H. pylori in regulating phenotypes linked to Sting deficiency within our murine model systems. Murine gastroid organoid monolayers were infected ex vivo with H. pylori strains J166 or PMSS1 for 24 hours and were used to analyze Sting alone (UI), 2ʹ3'-cGAMP, and/or H. pylori strain J166 or H. pylori gDNA. (d) RIG-I+ or parental cells were challenged with PBS alone (UI), 3p-hpRNA with or without H. pylori strain J166 or H. pylori gDNA. (e) STING+ or parental cells were challenged with PBS alone (UI), 2ʹ3'-cGAMP, and/or H. pylori strain J166 in the presence of increasing concentrations of Ruxolitinib. (f) RIG-I+ or parental cells were challenged with PBS alone (UI), 3p-hpRNA and/or H. pylori strain J166 in the presence of increasing concentrations of Ruxolitinib. STING and RIG-I activation was assessed and data are shown as fold activation relative to uninfected control. Experiments were performed in triplicate, and samples were run in duplicate within each experiment. ANOVA with Bonferroni correction was used to determine statistical significance between groups. (g) STING+ or parental cells were challenged with PBS alone (UI), 2ʹ3'-cGAMP with or without H. pylori strain J166 for 6 and 24 hours and were then assessed for IRF3 expression and activation by Western blot analysis. For Western blot analysis, conditions were tested at least 3 times and student's t-tests were used to determine statistical significance between groups. *p < .05, **p < .01, ***p < .001, ****p < .0001, ns = not significant. downstream pathways via RT-PCR. IRF3-dependent type I interferon stimulated genes Mx1 and Cxcl10 were significantly upregulated in wild-type murine gastric organoids following co-culture with 2ʹ3'-cGAMP, but expression was significantly reduced in samples co-infected with H. pylori and 2ʹ3'-cGAMP, and no changes were observed in Sting −/− monolayers (Figure 7(a-b)). Collectively, these data illustrate the differences between in vivo and ex vivo models in terms of the microenvironment, as the RNA-seq data generated from mouse tissue and RT-PCR analysis from mouse and human gastric organoids show differential regulation of Mx1 and Cxcl10. We speculate that in isolated organoid models, which lack immune components and other elements of the gastric microenvironment, differential responses in Stingdependent regulation of Mx1 and Cxcl10 may occur.
We next sought to further refine the identification of mediators of Sting suppression in response to H. pylori by filtering the overall differentially expressed gene lists to identify Sting-dependent genes. Using the comparison 1 list (Figure 6(b); Supplemental Table 2), any differentially expressed gene that was also identified in comparison 2 was removed ( Figure 6(b); Supplemental Table 3), leaving only 92 genes whose differential expression depended on the presence of Sting during H. pylori infection (Figure 6 Table 2). The resulting heatmap identified a target that has previously been shown to directly suppress Sting in mice, tripartite motif-containing 30a (Trim30a, Figure 8(b)). TRIM family proteins regulate critical cellular processes such as innate immunity, transcription, and autophagy 58-60 and can serve as effectors of innate immunity in response to signaling by cytokines such as IFN and tumor necrosis factor alpha (TNFα) and via pattern recognition receptors (PRRs) such as TLR, RIG-I, and STING. [60][61][62][63] Altered levels of Trim30a expression were subsequently validated by RT-PCR and significantly higher levels of Trim30a were observed in infected wild-type mice compared to Sting −/− mice (Figure 8(c)). To more precisely define the topography of Trim30a protein expression and clarify localization of the Trim30a in the gastric niche, immunohistochemistry was performed on the murine gastric tissue. Positive staining was observed predominantly in immune cells, such as polymorphonuclear leukocytes (PMNs), and some epithelial cells (Figure 8(d)). When immunohistochemistry was quantitated, H. pylori infection induced significantly higher levels of Trim30a expression compared to uninfected controls in both wild-type and Sting −/− mice. However, infected Sting −/− mice had significantly lower levels of Trim30a compared to infected wild-type mice (Figure 8(d)), similar to the RNA-seq and RT-PCR data.
In TLR-and STING-mediated signaling, TRIM30a serves as an important negative feedback regulator that controls excessive inflammatory responses via suppression of type I IFNs production and is expressed in a variety of cell types. [63][64][65] Therefore, we next sought to further investigate ex vivo and in vitro Trim30a expression by H. pylori in a cell-specific manner. To examine Trim30a protein expression within gastric epithelial cells, Western blot analysis was performed on co-culture lysates from murine gastric organoid monolayers infected ex vivo with H. pylori strains J166 and PMSS1 for 24 hours. Significantly increased levels of Trim30a expression were present in both H. pylori-infected wild-type and Sting −/− organoids compared to uninfected cells where no expression was observed (Figure 9(a)). Comparable Trim30a expression patterns were also demonstrated when examined by immunofluorescence in wild-type and Sting −/− organoids following a 24-hour co-culture with H. pylori (Supplemental Figure 6).
Bone marrow derived dendritic cells (BMDC) are also an important source of Trim30a, 63,64 thus, we next sought to determine if this pathway also functioned similarly within the context of H. pyloriinduced disease. Therefore, total mRNA was extracted from BMDCs isolated from wild-type or Sting −/− mice treated with or without H. pylori to quantify expression levels of Trim30a. H. pylori was determined by quantifying levels of phosphorylated IRF3 (p-IRF3, arrow) in gastric organoid co-cultures. Representative Western blots and densitometric analyses normalizing levels of pSTING and pIRF3 to total levels are shown at each time point. (c) RT-PCR analysis of MX1 and CXCL10 transcript levels are represented as relative gene expression levels normalized to levels of GAPDH gene expression. (d) Induction of autophagy was determined by quantifying levels of LC3-II. Representative Western blots and densitometric analyses normalizing levels of LC3-II to GAPDH are shown at each time point. In each experiment, conditions were tested at least 3 times and student's t-tests were used to determine statistical significance between groups. *p < .05, **p < .01, ***p < .001, ns = not significant.  (Figure 9(b)). Western blot analysis on BMDC protein lysates co-cultured with and without H. pylori revealed similar findings, with lower Trim30a protein levels observed in infected Sting −/− BMDCs compared to wild-type BMDCs (Figure 9(c)).
Collectively, in more physiologically relevant in vivo murine models, RT-PCR and immunohistochemistry demonstrate that H. pylori infection significantly increases Trim30a mRNA and protein expression and this response is partially attenuated in Sting-deficient mice. In ex vivo BMDCs, we observe similar findings, whereby RT-PCR and Western blot analysis demonstrate that H. pylori significantly increased Trim30a mRNA and protein expression and this response is partially attenuated in BMDCs isolated from Sting-deficient mice. It is important to consider however, that isolated peripheral BMDCs may function differently than gastric tissue DCs, which have not been directly assessed in this ex vivo model system. Since, in murine gastric organoids, Western blot analysis demonstrated that H. pylori significantly increased Trim30a expression, but no differences were observed between gastric organoids isolated from wild-type and Sting-deficient animals. These data illustrate the differences between in vivo and ex vivo models, and we speculate that the differences in Trim30a expression between these models is likely governed by the fact that the murine gastric organoids lack immune components and other elements of the gastric microenvironment that are present in the in vivo milieu.
The mouse specific Trim30a shares greatest homology with specific human TRIMs including TRIM5, TRIM6, and TRIM22 (Figure 10(a)). To extend our findings into human patients, we utilized tissues from a gastric cancer patient cohort to probe for TRIM30a human ortholog expression by RT-PCR. Expression of TRIM6 and TRIM22, but not TRIM5, was significantly increased in patient samples that harbored inflammation or cancer (Figure 10(b-c); Supplemental Figure 7). Another human TRIM homolog, TRIM29, has been directly implicated in STING modulation as well as gastric cancer outcomes; 66-68 thus, we also analyzed expression levels of TRIM29 within the patient cohort and demonstrated significantly higher levels of expression in patient samples that harbored inflammation or cancer (Figure 10(d)). These results raise the possibility that TRIMs represent targets induced by H. pylori infection, that can suppress STING activation and promote proinflammatory and pro-tumorigenic responses in vivo.

Discussion
Our laboratory has previously demonstrated that H. pylori can translocate DNA into host cells activating TLR9 18,31 but the role of other innate immune nucleic acid sensors during H. pylori infection has remained undefined. We now demonstrate using in vivo, ex vivo, and in vitro data that both STING and RIG-I-associated signaling was suppressed in the presence of viable H. pylori. One possible host mechanism for this phenomenon uncovered in our RNA-seq analysis is upregulation of TRIM proteins, which are known innate immune modulators. One such TRIM that was only upregulated in the presence of Sting and H. pylori was Trim30a, a known STING suppressor. 63 TRIM30a inhibits NF-κB activation induced by TLR signaling, including TLR9, via a K48linked ubiquitination mechanism that degrades Table 2 and TAB. However, NF-κB activation is required for initial upregulation of TRIM30a colonization in infected mice. (b) Acute inflammation was assessed and scored in the antrum and corpus of C57BL/6 wild-type or Sting −/− mice infected with or without H. pylori by a pathologist blinded to treatment groups. Histologic parameters were scored according to the Sydney System. 54 64 This suggests that TRIM30a may be initially induced by H. pylori infection through activation of TLR9 which can then act as a negative regulator of both TLR9 and STING to dampen the subsequent immune response to H. pylori. However, to directly assess the role of TRIM30a in the negative regulation of STING, experimental reduction of TRIM30a is needed to more directly assess the effects on both STING and IRF3 activation.
Other Trim genes revealed to be upregulated by H. pylori (Supplemental Tables 1 and 2) that can modulate innate immune suppression include Trim40, which targets the downstream RIG-I regulator MAVS for K48-linked ubiquitination 62 and Trim10, which can suppresses IFN/JAK/STAT signaling pathway through blocking the interaction between IFNAR1 and TYK2 to negatively regulate type I IFN signal transduction. 69 Our in vitro and ex vivo work also demonstrated that H. pylori could directly mediate innate immune signaling via suppression of IRF3 activation.
Although, we have not directly assessed the role of the cag type IV secretion system in this study, it has been well established that the cag T4SS translocates the oncoprotein CagA as well as microbial DNA, which can be sensed by innate immune receptors. 18,31 Another study by Song et al. 11 demonstrated that H. pylori infection increases Sting expression and p-Irf3 expression by immunohistochemistry and Ifnb gene expression in murine gastric tissue. Although the authors also used C57BL/6 mice, they used a strain of H. pylori (SS1) that does not have a functional cag type IV secretion system (T4SS) and thus cannot deliver the proinflammatory protein CagA or DNA via the T4SS. We speculate that the lack of a functional T4SS may account of for the differences observed in this model, compared to the current models.
This complex system of innate immune suppression and activation builds on our previous work focused on the duality of TLR9 signaling during H. pylori infection and suggests that DNA translocation, induction of TRIM proteins, and suppression of IRF3 activation may be yet another component of a finely tuned rheostat that H. pylori utilizes to regulate the initial innate immune response, and ultimately drive long term activation of inflammatory pathways such as those promoted by increased Th17 activation.
In conclusion, this study demonstrates that H. pylori actively suppresses innate nucleic acid sensors STING and RIG-I potentially via downregulation of IRF3 activation and induction of TRIM proteins. Additionally, loss of Sting augments acute inflammatory responses to H. pylori within the in vivo gastric niche. This work lays the foundation for further exploration into the role of H. pyloriinduced TRIMs in human hosts and suggest that manipulation of TRIMs may represent a novel strategy to prevent or treat pathologic outcomes induced by H. pylori infection.

Helicobacter pylori
Wild-type H. pylori cag + strains J166, PMSS1, 18 H. pylori strains were analyzed for growth as previously described. 72 Briefly, overnight cultures were sub-cultured in a 96 well flatbottom plates and incubated in a microaerophilic chamber, as described above. Optical densities (OD) were recorded at 600 nm (BioTek) at 2, 4, 6, 8, 12, and 24 hours. The final OD value was normalized using uninfected media as a negative control.

Murine models of Sting deficiency
All animal studies were carried out in accordance with the recommendations in the Guide for the Care

Histopathology
A single pathologist (MBP), blinded to treatment groups, scored indices of inflammation and injury as described previously. 17,73 Specifically, the following variables were graded on a 0 to 3 scale (0, none; 1, mild; 2, moderate; 3, severe) in the gastric antrum and body: acute inflammation (polymorphonuclear cell infiltration) and chronic inflammation (mononuclear cell infiltration independent of lymphoid follicles); thus a maximum inflammation score of 12 was possible for each animal.
Anti-CD3 antibody #Ab16669 (Abcam; 1:250) and anti-TRIM30 antibody #NBP2-41087 (Novus Biologicals; 10 μg/ml) were used to stain gastric tissue sections by the Vanderbilt University Medical Center Translational Pathology Shared Resource (TPSR). A single pathologist (MBP), blinded to treatment groups, evaluated and scored all immunohistochemistry (IHC). Trim30a, MPO, CD3, CD68, CD45, and CD11c were evaluated by quantifying positive cells in 5 HPFs (400x) with the highest counts in each mouse. Sting and RigI staining were evaluated by assessing the percentage of positive epithelial cells and grading the intensity of staining in epithelial cells semi-quantitatively, as previously described. 74

pylori infection of murine gastric organoids downregulates IRF3-dependent type I interferon stimulated genes.
Murine gastric organoid monolayers were challenged with PBS alone (UI), STING agonist 2ʹ3'-cGAMP, and/or wild-type (wt) H. pylori strain J166 or PMSS1 at MOI 100:1 for 6 or 24 hours. RT-PCR analysis of (a) Mx1 and (b) Cxcl10 transcript levels was assessed in cocultured murine gastric organoids. Data are represented as relative gene expression levels normalized to levels of Gapdh gene expression. In each experiment, conditions were tested at least 3 times and student's t-tests were used to determine statistical significance between groups. *p < .05, **p < .01, ***p < .001, ****p < .0001. were then converted to 2D epithelial cell monolayers following Matrigel removal and 3D gastric organoids were plated on collagen-coated plates. Bone marrow-derived dendritic cells (BMDC) were generated from bone marrow obtained from femurs of wild-type and Sting −/− mice. Briefly, marrows were treated with red blood cell lysis buffer (KD Medical) and washed with PBS and recovered white blood cells were plated in advanced DMEM media (Gibco) supplemented with 20% FBS and 40 ng/mL each of GM-CSF and IL-4 (Peprotech) for 6 days at 37°C and 5% CO 2 for differentiation.

Ex vivo and in vitro infections
Primary 2D gastric organoid monolayers were co-cultured with or without 2ʹ3'-cGAMP (30 μg/ml) and H. pylori strains J166 or PMSS1 at a multiplicity of infection (MOI) of 100:1 for 6 or 24 hours. BMDCs were cocultured with wild-type H. pylori strain J166 at a multiplicity of infection (MOI) of 10:1 or ODN-1826 (6 μg/ml) for 6, 12, or 24 hours. RNA and protein were then isolated from coculture samples for RT-PCR and Western blot analysis, respectively.

STING reporter assay
HEK293 hSTING-R232 cells (STING+) and HEK293 Null1 (Parental) cells were seeded in 96well plates (Corning) at 50,000 cells per well in DMEM without antibiotics and challenged with either viable H. pylori (MOI 100:1), sterile PBS, and/or 2ʹ3-cGAMP (30 μg/ml) at 37°C with 5% CO 2 for 24 hours. Supernatants were then added to QUANTI-Blue™ solution (Invivogen) and plates were analyzed by spectrophotometer (Bitoek) at 650 nm. All experiments were performed in duplicate and repeated at least three times. Data are expressed as fold over uninfected control.

RIG-I reporter assay
HEK-Lucia RIG-I cells (RIG-I+), and HEK-Lucia Null cells (RIG-I Parental) cells were seeded in 96-well plates (Corning) at 50,000 cells per well in DMEM without antibiotics and challenged with either viable H. pylori (MOI 100:1), sterile PBS, and/or 3php-RNA (5000 ng/ml) at 37°C with 5% CO 2 for 24 hours. Supernatants were then added to QUANTI-Luc™ solution (Invivogen) and plates were analyzed by luminometer (Bitoek). Experiments were performed in triplicate, and samples were run in duplicate within each experiment. Data are expressed as fold over uninfected control.

Cell viability assay
The effect of H. pylori and/or agonists on reporter cell viability was assessed in STING+, RIG-I+, and their respective parental cells using the CellTiter-Blue assay (Promega), according to the manufacturer's instructions. In brief, following co-culture, STING or RIG-I reporter cells were washed with PBS containing gentamycin (250 μg/ml) and hygromycin (500 μg/ml), followed by incubation with DMEM media containing gentamycin (250 μg/ml) and hygromycin (500 μg/ ml). After a 30-minute incubation at 37°C to eliminate residual viable H. pylori, CellTiter-Blue reagent was added. Samples were incubated for 1 hour at 37°C and fluorescence was measured (485 Ex /516 Em ) using a fluorescent imaging plate reader (Biotek).

Real-time PCR
RNA was extracted using the RNAeasy Mini Kit (Qiagen) for all sample types, according to the manufacturer's instructions. cDNA was synthesized  Representative Western blots and densitometric analysis normalizing levels of Trim30a to Gapdh. In each experiment, conditions were tested at least 3 times and student's t-tests were used to determine statistical significance between groups. *p < .05, **p < .01, ns = not significant. ## p < .01, ### p < .001 #### p < .0001 compared to untreated.
using High-Capacity cDNA Reverse Transcription Kit (ThermoFisher) and quantitative real-time PCR was performed using Power SYBR Green Master Mix (ThermoFisher) for human samples and TaqMan™ Universal Master Mix II (ThermoFisher) for murine samples, with gene-specific primers (Table 3).

Western blot analysis
AGS cells, human, and murine organoids co-cultured with H. pylori were and protein lysates were separated using 6% (AGS cells) or 10% (organoids) SDS PAGE mini gels, transferred to PVDF membranes, and membranes were blocked with BSA or milk as ). An anti-rabbit or antimouse HRP-conjugated secondary antibody (Promega; 1:10000) was then incubated with membranes for 1 hour. Protein intensities were quantified using ImageJ software (NIH).

Trim30a immunofluorescence staining
Monolayers of primary gastric epithelial cells derived from C57BL/6 wild-type and Sting −/− mice were infected for 24 hours with H. pylori strains J166 or PMSS1. After infection, monolayers were subjected to immunofluorescence staining as previously described. 17,50 Briefly, cells were fixed with 10% neutral-buffered formalin (Azer Scientific), permeabilized with Triton X-100 (Promega), and then blocked with Dako Protein Block Solution (Agilent) for 1 hour. Samples were incubated with anti-TRIM30 rabbit antibody #NBP2-41087 (Novus Biologicals; 1:50) overnight at 4°C before detection with Alexa-fluor secondary antibody (Invitrogen). Nuclei were detected using Hoescht (Invitrogen). Slides were mounted using ProLong Glass (Invitrogen), and images were acquired in an Olympus FV-1000 confocal microscope. Experiments were performed in part through the use of the Vanderbilt Cell Imaging Shared Resource (CISR).

RNA sequencing and analysis
Total RNA from wild-type C57BL/6 and Sting −/− mice was processed using a NEBNext® Ultra™ II RNA Library Prep sample prep kit following the manufacturer's instructions (New England Biosciences) and evaluated on a Qubit 3.0 fluorometer and an Agilent 2100 Bioanalyzer to quantitate concentration and fragment size distribution prior to sequencing using the NovaSeq 6000 sequencer (Illumina). Sequencing was performed using a S4 flow cell with a PE 150 kit. Individually barcoded libraries were then pooled at an equal molar ratio and sequenced at 2 × 150 bp/read. Approximately 48 million paired-end sequence reads per sample (mean ± SD = 48.3856 ± 7.655 million; n = 34) were generated. Sample quality was assessed via FastQC software. The data were analyzed using the Dragen Software Version: 3.6.3 aligning the data to the mm10 reference genome. The Vanderbilt Technologies for Advanced Genomics (VANTAGE) core facility prepared the RNA library, assessed library quality, and performed sequencing. Differential expression analyses were performed at baseline (uninfected WT C57BL/6 versus Sting −/− ) and based on the following 2 major comparisons: wild-type C57BL/6 infected versus wild-type C57BL/6 Brucella broth control, and Sting −/− infected versus Sting −/− Brucella broth control. Filters used to identify differential expression were an adjusted p-value <0.05 and an absolute log2 fold change >1. Venn diagrams were created (Venny 2.1 software) using these comparisons. Ingenuity Pathway Analysis software (Qiagen) was used to link differentially expressed genes in the dataset to particular biological functions and pathways.

Human clinical specimens
Snap frozen de-identified human gastric tissue samples were acquired from the Cooperative Human Tissue Network (CHTN). Normal gastric tissue, or gastric tissues harboring either gastritis alone or gastric adenocarcinoma were disrupted and homogenized using ZR BashingBead Lysis tubes (Zymo Research) prior to RNA extraction. The protocol was approved by the Vanderbilt University Medical Center Institutional Review Board (#210729).

Statistics
The student's t-test was used for two group comparisons, while one-way analysis of variance (ANOVA) with Bonferroni correction was used for multiple group comparisons. Data were plotted and analyzed using Prism 6.0 (GraphPad). Statistical significance was set at a two-tailed p-value of <0.05. In all figures, means ± standard errors of the mean are shown.

Ethics of experimentation
All animal and human studies were conducted in accordance with the Declaration of Helsinki principles and have been approved by the Vanderbilt University Medical Center Institutional Animal Care and Use Committee (IACUC) and the Vanderbilt University Medical Center Institutional Review Board (IRB), respectively. Human gastric tissue samples used for RT-PCR were acquired from the Cooperative Human Tissue Network and these analyses were approved under IRB #210729 under the category of a nonhuman subject study.

Disclosure statement
No potential conflict of interest was reported by the author(s).